1232 | Metallomics, 2018, 10, 1232--1244 This journal is©The Royal Society of Chemistry 2018

Cite this:Metallomics, 2018,

10, 1232

Short oligopeptides with three cysteine residuesas models of sulphur-rich Cu(I)- and Hg(II)-bindingsites in proteins†

Edit Mesterhazy,ab Colette Lebrun,b Serge Crouzy,c Attila Jancso *b andPascale Delangle *a

The essential Cu(I) and the toxic Hg(II) ions possess similar coordination properties, and therefore, similar

cysteine rich proteins participate in the control of their intracellular concentration. In this work we

present the metal binding properties of linear and cyclic model peptides incorporating the three-

cysteine motifs, CxCxxC or CxCxC, found in metallothioneins. Cu(I) binding to the series of peptides at

physiological pH revealed to be rather complicated, with the formation of mixtures of polymetallic

species. In contrast, the Hg(II) complexes display well-defined structures with spectroscopic features

characteristic for a HgS2 and HgS3 coordination mode at pH = 2.0 and 7.4, respectively. Stability data

reflect a ca. 20 orders of magnitude larger affinity of the peptides for Hg(II) (logbpH7.4HgP E 41) than for

Cu(I) (logbpH7.4CuP E 18). The different behaviour with the two metal ions demonstrates that the use of

Hg(II) as a probe for Cu(I), coordinated by thiolate ligands in water, may not always be fully appropriate.

Significance to metallomicsMercury(II) and copper(I) detoxification in cells involves similar sulphur-rich proteins, including metallothioneins. Short model peptides are used here to studythe molecular interactions of these two soft cations with sequences containing three cysteine moieties found in these proteins. We demonstrate that thebinding of Hg(II) and Cu(I) with these representative cysteine-rich sequences involves different molecular mechanisms, with the formation of variouscoordination complexes. Hence, the use of Hg(II) as a probe for Cu(I) coordination with sulphur-rich peptides or proteins in physiological conditions isdemonstrated here to be not fully appropriate.

Introduction

Many metal ions, including Fe(II/III), Cu(I/II) or Zn(II), are essentialmicronutrients. Since both their deficiency and excess areharmful for the cells, the optimal concentration of these ionsis controlled by regulation systems involving lots of proteins.Other metal ions, like Hg(II) and Cd(II), are toxic without anyadvantageous effects. The high affinity of these soft metal ionsfor soft sulphur bases, found in proteins and small ligands,such as glutathione, is one of the main factors leading to theirtoxicity. Indeed, interactions between these toxic metal ions

and thiol-containing molecules induce misfolded proteinstructures or perturbation in the redox balance of the cells.

As a consequence of the similar coordination propertiesof the essential Cu(I) and the toxic Hg(II) ions, the cellularregulation and protection systems for controlling or reducingthe concentration of these ions involve proteins with sulphur-richmetal coordination sites, usually dominated by Cys-thiolate donorgroups. A varying number of Cys residues may allow the bindingof Cu(I) or Hg(II) in isolated monometallic coordination sites orin polymetallic clusters. The strong preference of Hg(II) for a bis-thiolate coordination environment is reflected by the publishedstructures of several elements of the bacterial mercury resistancesystem, MerP,1 MerT,2 the N-terminal domain of MerA3 or theHg(II)-bound form of the copper chaperone Atx1.4 Nevertheless,there are also examples of structures with more Hg(II)–thiolatebonds, e.g. the distorted tetrahedral tetrathiolate Hg(II)-centreof rubredoxin,5 or the tri- or tetra-coordinated metal ionenvironments in Hg(II)-bridged protein dimers, like thoseobserved in the MerR metalloregulatory protein6 or in the human

a INAC/SYMMES/Universite Grenoble Alpes, CEA, CNRS, 38000 Grenoble, France.

E-mail: pascale.delangle@cea.frb Department of Inorganic and Analytical Chemistry, University of Szeged,

Dom ter 7, Szeged H-6720, Hungary. E-mail: jancso@chem.u-szeged.huc BIG/LCBM/Universite Grenoble Alpes, CEA, CNRS, (UMR 5249), 38000 Grenoble,

France

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c8mt00113h

Received 21st May 2018,Accepted 13th July 2018

DOI: 10.1039/c8mt00113h

rsc.li/metallomics

Metallomics

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This journal is©The Royal Society of Chemistry 2018 Metallomics, 2018, 10, 1232--1244 | 1233

copper chaperone HAH1.7,8 Cu(I) can also accommodate a linearCuS2-type coordination mode, as indicated by structural data onthe copper efflux regulator CueR9 or the metal binding domainsof the copper transporter protein ATP7B.10 Trigonal CuS3 centreswere proposed in the binuclear Cu(I)–thiolate core of the repressorprotein CopY11 and in one of the suggested metal-bridged dimericforms of the copper chaperone CopZ,12 whereas the Cu(I)-bridgedHAH1 dimer7 was shown also to include a fourth, weakly bound,thiolate in a pseudotetrahedral environment.

Di-, tri- or tetrathiolate coordination modes are also typicalin polymetallic clusters. Metallothioneins (MTs) are small,cysteine-rich proteins, found in all kingdoms of life fromprokaryotes to mammals. Several isoforms of MTs evolvedwith high diversity of amino acid sequences and, pursuant tothis, with different metal ion preferences, 3D structures andproposed biological functions.13,14 A novel approach for theclassification of such a big family of proteins was developed bythe groups of Capdevila and Atrian based on the metal bindingfeatures of MTs, i.e. on the criterion whether the MT bindsZn(II) either in homo or in heterometallic form, or only Cu(I)under physiological conditions.15,16 Trigonally coordinatedCu(I) ions strongly dominate in Cu(I)-loaded MTs displayingalso less abundant Cu(I)S2 centres.17,18 The only publishedCu(I)-loaded MT crystal structure, a truncated form of theCu(I)–MT of yeast, bears a Cu(I)8S10 core with six tri- and twodi-coordinated metal ions where all Cys residues bridge two orthree Cu(I) centres.19 It was also suggested that a Cu(I)4S6

cluster could form in yeast MTs under low Cu(I) availability.20

Indeed, the latter core contains only tri-coordinated Cu(I) and iswidely used by nature in many other types of metalloproteins,21

e.g. in the C-terminal domain of the membrane copper transporterCtr122 or in various Cu(I)-activated transcription factors.23,24

In contrast, Hg(II)–thiolate clusters in MTs were found to displaya complicated mixture of di-, tri- and tetra-coordinated Hg(II)centres, with strong dependence on the applied conditions(pH, temperature, Hg(II) concentration, counter ions, etc.).25,26

An interesting strategy to better understand the individualcontributions of specific short sequences to the overall affinityof proteins for metal ions is to design and study modelpeptides. This approach could eventually lead to the identifi-cation of relevant peptide-based motifs for heavy metal chelation.Some of the above described metal-binding sites have beenprobed by using oligopeptides encompassing the relevant metalbinding sequences of the modelled proteins. The metal-bindingfeatures of the metalloregulator CueR and the metallochaperoneAtx1, both encompassing 2 cysteines in their metal-bindingdomain, with different sequences between the Cys residues, wereinvestigated via oligopeptide models of the relevant metal-bindingloops.27–33 A general finding was that the linear unstructuredligands, while displaying high affinity and interesting metal ionselectivity for Hg(II) or Cu(I) chelation, could not fully reproducethe efficiency of the modelled proteins.28,31,33 Cyclisation of thelinear Atx1 model, resulting in a more rigid skeleton with pre-oriented Cys-sidechains, was an important step forward leadingto notably increased metal binding affinities.27,28 The morechallenging imitation of the HgS3 type metal coordination sites

was successfully achieved by different approaches. Pecoraro et al.successfully applied single oligopeptide chain three-helixbundles34 and three-stranded coiled coils35 to settle three Cysresidues into optimal position for accommodating the Hg(II)ion in a tri-coordinate fashion under slightly alkaline conditions.These tris-thiolate bound species were in a pH-dependent equili-brium with HgS2 type complexes, characterized by apparent pKa

values Z7.34,35 Using a different strategy, tripodal pseudopeptideligands were designed with cysteine or D-penicillamine (D-Pen)moieties grafted onto a nitrilotriacetic acid scaffold.36,37 Theseconstructs were able to stabilize the HgS3 coordination mode in abroad pH-range, starting even at pH B 5.5 with one of the D-Penligands.36 Interestingly, the behaviour of the same ligands inbinding Cu(I) was less straightforward. Depending on thebulkiness and hydrophobicity of the arms attached to thetripodal template, the formation of mononuclear and (Cu2Lig)x

type complexes with (Cu2S3)x cores was observed with highstabilities.38,39 In a recently published article, the Hg(II) bindingof the cyclic peptide P3C incorporating a three Cys variant of themetal binding loop of Atx1 was discussed.40 This cyclic solventaccessible peptide was considered as a predisposed structurethat could pre-orient the Cys-sidechains for an easy accommo-dation of metal ions in a tri-coordinate fashion. The resultsconfirmed the formation of a mononuclear Hg(II) complex withproperties indicative of an HgS3 coordination.

Peptide scaffolds similar to P3C and comprising two typicalmetal binding motifs found in metallothioneins, namely theCxCxxC and CxCxC sequences,41–43 are investigated in thepresent paper. Their binding properties for the two softmetal ions Cu(I) and Hg(II), exhibiting the largest affinities formetallothioneins, are studied and discussed in relation to theirbinding by detoxification systems.26,44–46

Experimental sectionMaterials

Materials and solvents were purchased from Sigma Aldrich,Fluka, Acros Organics and used without further purification.Amino acid building blocks, resins and coupling agents werepurchased from NovaBiochem. For aqueous solutions, ultrapurelaboratory grade MilliQ water was used (resistivity 18 MO cm).

Abbreviations

Ac2O: acetic anhydride; AcN: acetonitrile; BCS: bathocuproine-disulfonate; DCM: dichloromethane; DIEA: N,N-diisopropyl-ethylamine; DTNB: 5,50-dithio-bis-(2-nitrobenzoic acid); EDT:ethanedithiol; Et2O: diethyl ether; Fmoc: 9-fluoroenylmethoxy-carbonyl; HBTU: 2-(1H-benzotriazol-1-yl)-1,1,3,3-tetramethyl-uronium hexafluorophosphate; HOBt: hydroxybenzotriazole;MeOH: methanol; NMP: N-methyl-2-pyrrolidone; TFA: trifluoroacetic acid; TIS: triisopropylsilane.

Peptide synthesis and purification

Linear peptides. Ac-GCTCSCGSRP-NH2 (1L) andAc-GTCSCGCSRP-NH2 (2L) peptides with acetyl and amide

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protection at their two termini were synthesized by standardsolid phase peptide synthesis following the Fmoc procedure.Rink Amide AM resin (200–400 mesh, 0.67 mmol g�1) was usedas solid support. The average coupling mixture consisted of4 equiv. of protected amino acid building blocks, 4 equiv. ofHOBt, 4 equiv. of HBTU and 8 equiv. of DIEA dissolved in NMP.The reaction time was 1 h. The noncomplete coupling wasmonitored by Kaiser’s test.47 To avoid the formation of deletedproducts, the acetylation of the unreacted amino groups wasperformed in a mixture of 10% Ac2O and 10% DIEA in DCMafter each coupling reaction. The Fmoc deprotection wasachieved by 20% piperidine solution in NMP. The cleavage ofthe peptide from the resin and the removal of the sidechainprotecting groups were performed in one step in the mixture ofTFA/EDT/H2O/phenol/TIS (92/2.5/2.5/2/1 V/V%) for 4 h. Afterevaporation, the residue was precipitated in ice cold Et2O andcentrifuged (6000 rpm, 15 min).

Cyclic peptides. For c(SRPGCTCSCG) (1C), c(CSRPGTCSCG)(2C) and c(CSRPGATCSCG) (3C), linear precursors were firstassembled on preloaded H-Gly-2-Cl-trityl resin (0.54 mmol g�1)following the same method as described above. The protectedprecursor was cleaved from the resin by treatment with 2% TFA inDCM for 2� 5 min. The solution was flushed into 20% pyridine inMeOH. After concentration, the residue was precipitated in icecold Et2O. The protected linear precursor (0.5 mM in DCM) wasreacted with 3 equiv. of PyBOP and 4 equiv. of DIEA. After 2 hreaction time, the DCM was removed by evaporation and theresidue was precipitated in ice cold Et2O. The sidechain protectinggroups were removed in TFA/EDT/H2O/TIS (90/5/2.5/2.5 V/V%) for2 h. After evaporation, the remaining solution was precipitated inice cold Et2O.

The crude products were purified by reversed phase HPLCon a Shimadzu LC-20 instrument equipped with a PhenomenexSynergi 4 mm fusion-RP 80 Å semi-preparative column. H2O andacetonitrile with 0.1% TFA were used as eluents. The purity waschecked by analytical HPLC and ESI-MS. Analytical RP-HPLCwas performed using an analytical column (Chromoliths

performance RP-18e 100-4.6 mm) at 1 mL min�1 flow rate(gradient 0–60% of AcN in 30 min). The synthesis of P3C hasbeen published earlier.40 For the other peptides analyticalHPLC and ESI-MS characterization is available in Table S1and Fig. S1 in the ESI.†

Sample preparation for physicochemical studies

The cysteine residues of the peptides and Cu(I) are sensitive tooxidation; therefore all sample preparations and manipulationswere performed in a glove box, under an argon atmosphere.Fresh solution was made for each experiment in the appro-priate buffer. 10% acetonitrile in volume was added to the Cu(I)containing samples to overcome Cu(I) disproportion in water.48

The final peptide concentration was determined by Ellman’sprocedure, where DTNB reacts with the free thiols formingTNB� (2-nitro-5-thiobenzoate) with e412 = 14 150 M�1 cm�1.49,50

Cu(I) solution was made from Cu(CH3CN)4PF6 salt in acetonitrile.The final concentration was determined by using BCS forminga stable Cu(BCS)2 complex with e483nm = 13 300 M�1 cm�1.51

A precise weight of high purity HgCl2 was used to prepare Hg(II)stock solution in water (B3 mM).

UV-Vis absorption and circular dichroism (CD) measurements

UV-Vis spectra were recorded on a Varian Cary 50 spectro-photometer equipped with optical fibers connected to an externalcell holder in the glovebox. CD spectra were acquired withan Applied Photophysics Chirascan photometer. (1S)-(+)-10-Camphorsulfonic acid served as the calibration compound forthe instrument. Spectra were recorded in the 320–190 nmwavelength range with 1 nm bandwidth and 2 s dwell timeper point. For each sample 3 parallel spectra were recorded andthe average of these spectra was smoothed by the Savitzky–Golay method with a ‘‘window size’’ 7. CD spectra arereported in molar ellipticity ([Y] in units of deg cm2 dmol�1).[Y] = yobs/(10lc), where yobs is the observed ellipticity in milli-degrees, l is the optical path length in cm and c is the peptideconcentration in mol dm�3.

With Cu(I), titrations were performed in individually preparedsamples because of the slow complex formation. 2.5 mL volumesof peptide solution (20–30 mM) in phosphate buffer (20 mM,pH = 7.4) were transferred into UV cells (1.0 cm path length) andthen different (0.0–3.0) equivalents of Cu(I) were added to thesesamples. The samples were equilibrated for at least 2 hours.To ensure the thermodynamic equilibrium was reached, theabsorbance was measured regularly. In the case of Hg(II)titrations, 2.5 mL of peptide solution (20–30 mM) in phosphatebuffer (20 mM, pH = 7.4) was transferred into a UV cell(1.0 cm path length) and then aliquots of the Hg(II) ion solution(B3 mM) were gradually added from 0.0 to 3.0 equivalents. pHtitrations of the Hg(II) complexes were performed in pure waterin the pH range of 2–11 by adding aliquots of 0.1 M KOH. ThepH was measured using a Metrohm 702 SM Titrino equippedwith a Mettler Toledo InLabs Micro electrode. pKa values forthe observed deprotonation processes were obtained by fittingthe data by using the SPECFIT computer program.52–55

ESI-MS experiments

Mass spectra were acquired on a LXQ-linear ion trap (THERMOScientific) instrument equipped with an electrospray ionsource. Electrospray full scan spectra in the range of m/z =50–2000 amu were obtained by infusion through a fused silicatubing at a flow rate of 2–10 mL min�1. The solutions wereanalysed in negative and positive ion modes. The LXQ calibra-tion (m/z = 50–2000) was achieved according to the standardcalibration procedure from the manufacturer (mixture of caf-feine, MRFA and Ultramark 1621). The temperature of theheated capillary for the LXQ was set in the range of 200–250 1C, the ion-spray voltage was in the range of 3–6 kV andthe injection time was 5–200 ms. The ligand solution (100 mM)was prepared in ammonium acetate buffer (20 mM, pH = 6.9)and aliquots of the appropriate metal ion were then added.

Determination of apparent stabilities of the metal ion complexes

The apparent stability constants at a given pH were determinedby UV-Vis titration in the presence of a competitor. The Cu(I)

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complexes were prepared by adding 0.9 equivalents of Cu(I)to three samples of each peptide (50 mM) in phosphate buffer(20 mM, pH = 7.4)/acetonitrile (9/1 V/V). Then 4, 8 and 12equivalents (with respect to Cu(I) concentration) of BCS wereadded, respectively. The samples were equilibrated until theabsorbance stabilized. Stability constants were calculated con-sidering the formation of the CuP mononuclear complexesat the beginning of the titration according to the followingequation:

bpH7:411 ¼ ½CuP�½Cu�½P�

where CuP and P represent the complexed and free peptide atthe pH of the studies, independent of the protonation state ofthe ligand, and Cu stands for the free Cu(I) ions.

The stabilities of the Hg(II)–peptide complex were determinedat pH = 2.0 by using iodide (I�) as a competitor. Stability constantsof the HgIn (n = 1–4) complexes were taken from the literature56

and recalculated for the conditions of the experiments (pH = 2.0,I = 0.1 M NaClO4) by applying the SIT model.57,58 The molarabsorption spectra of the four Hg(II)–I� complexes were deter-mined by titrating a Hg(II) solution by the continuous addition ofaliquots of I� solutions (0.01, 0.1 and 0.5 M KI). The recordedspectra were fitted with SPECFIT52–55 by fixing the stability con-stants of the HgIn complexes. The peptide solutions were preparedin water (30 mM, I = 0.1 M NaClO4) and the pH was adjusted by a1.0 M HCl solution. Samples for the competition experimentswere prepared by adding 1.0 equivalent of Hg(II) to the peptidesolutions. They were then titrated by the I� solutions by thesame procedure and under the same conditions as describedabove. The pH of samples was under control throughout theexperiments. SPECFIT52–55 was used for the evaluation of spectra.The molar absorption spectra and the stabilities of the HgIn

complexes were fixed in the fitting procedure, allowing one tocalculate the apparent stability of the HgP binary and HgPI ternarycomplexes (see the equations below), as well as their molarabsorption spectra.

bpH2:0HgP ¼

½HgP�½Hg�½P�

bpH2:0HgPI ¼

½HgPI�½Hg�½P�½I�

In the equations above, HgP (or HgPI) and P represent thecomplexed and free peptide at the pH of the studies, indepen-dent of the protonation state of the ligand, and Hg and I standfor the free Hg(II) and I� ions, respectively.

Determination of acid dissociation constants (pKa)

Deprotonation processes of one of the linear peptide ligands,1L, were followed by pH-potentiometric titrations in aqueoussolution (T = 298.0 � 0.1 K, cligand = 1.0 � 10�3 M, I = 0.1 MNaClO4) following a protocol described earlier.29 An automatictitration set, including a PC-controlled Dosimat 665 (Metrohm)autoburette and an Orion 710A precision digital pH-meter

equipped with an Orion 8103BNUWP Ross Ultra semi micropH electrode, was used to carry out the experiments. Argonatmosphere in the titration cell was applied in order to preventthe oxidation of the ligand. The data obtained in two paralleltitrations were fitted using the PSEQUAD computer program59

based on the following general equilibrium process with therelated equilibrium formation constants allowing the calculationof the pKa values for the individual deprotonation steps:

qH + rL " HqLr

bHqLr¼

HqLr

� �

½H�q½L�r

where L denotes the non-protonated peptide and H the protons(charges are omitted for simplicity). Consequently, the compo-sition of the neutral fully protonated peptide is H3L. These pKa

values were used in the calculation of the thermodynamicstability constants for the HgL and HgHL complexes from theapparent stabilities obtained at pH = 2.0. Details of thesecalculations are found in the ESI.† Note that ‘L’ for the ligandsis used in the text where the actual protonation states of theligands/complexes are taken into account in calculations or inthe description of species, whereas ‘P’ denotes the peptides ingeneral, independent of their protonation states, as in thecalculation of the apparent stabilities.

Modelling of the trithiolate mercury complexes

The six peptides were modelled in the apo and Hg(II)-boundform with a HgS3 trigonal coordination. In the absence ofstructures for all the peptides but P3C, initial coordinates werebuilt with the CHARMM program60 from standard values forthe internal bonds, angles and dihedrals of amino acids inproteins. For peptide P3C, initial coordinates were either generatedas for other peptides or taken from the solution NMR structure ofthe HgP3C complex.40

Model parameters. For the holo peptides, a bonded model ofthe mercury site was used. Molecular mechanics force fieldparameters around mercury, absent from the CHARMM forcefields, were deduced from ab initio DFT/B3LYP61–63 calculationson the model system Hg2+–(S–CH2–CH3)3

� of tri-coordinatedmercury using the Gaussian program.64 A total charge of �1 inthe singlet spin state and using the LanL2dz basis set was used.Bonded parameters were obtained by harmonic potential fittingof ab initio scans of the Hg–S distance, C–S–Hg angle andHg–S–S–S improper angle. In this trigonal model, the S–Hgdistance is 2.794 Å, the Cb–S–Hg angle is 1251 and the Hg–S–S–Simproper angle (calculated with Hg above the 3 sulfur atom planein the counter clockwise direction) is 7.81. Charges were obtainedfrom RESP calculations following DFT/B3LYP optimization.Calculated charges are: Hg: +0.968, S: �0.798, all C(bonded to2 hydrogens): 0.173, all C(bonded to 3 hydrogens): 0.069. van derWaals parameters were taken from the work of Sebesta et al.65 forHg: e = 1.955 kcal mol�1; R = 2.8 Å.

Molecular dynamics simulations. The geometries of the pep-tides were relaxed in vacuum by 1000-step energy minimizationusing the conjugated gradient method until the root-mean-square

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(RMS) energy gradient was lower than 0.1 kcal mol�1 �1.Then the peptides were simulated in implicit water using theimplicit solvent method EEF1 and the adapted extended atomCHARMM19_EEF1 force field.66 The systems were subjected to40 to 300 ns MD simulations (depending on the convergencespeed) using Langevin dynamics with a time step of 2 fs and adamping factor fb of 20 ps�1 applied on non-hydrogen atoms.All peptides were simulated in the apo (no Hg) and holo (with Hgbound) forms.

Results and discussionPeptide design and synthesis

The cyclic peptide P3C with three cysteine sidechains preorientedfor Hg(II) coordination was recently reported by our group.40

All spectroscopic data obtained for the HgP3C mononuclearcomplex point to a HgS3 coordination mode, which is stable overa large pH range (pH = 5–9).

Five new peptides (Scheme 1) were designed in this workto monitor the effects of introducing a CxCxC motif instead ofthe CxCxxC sequence found in P3C on the Cu(I) and Hg(II)complexation features. The positions of the three cysteines inthe sequence of the newly designed linear and cyclic peptideswere varied. An xPGx b turn inducing motif was introduced inthe sequences of the cyclic peptides to rigidify the scaffoldcontaining the metal-binding fragment CxCxC next to the turnin 1C or one amino acid apart in 2C.28,67–69 1L and 2L linearanalogues of the cyclic decapeptides 1C and 2C were synthesizedto determine the impact of the cyclisation and flexibility of thebackbone on the coordination properties. Finally, an additionalamino acid, namely alanine, was inserted in peptide 3C with theidea of promoting larger flexibility in the cysteine sidechainorientation.

Cu(I) complexes

One of the six investigated peptides, P3C, was previously demon-strated to stabilize the HgS3 coordination in a mononuclearcomplex.40 This raised the question whether a CuS3 geometrycould also be observed with the latter peptide and with the new3-Cys containing peptides presented in this paper. Cu(I) com-plexation of the ligands was studied by UV-Vis and CD spectro-scopy and ESI-MS. All the six peptides display a rather similarbehaviour; therefore studies with peptide 1C are presented hereto illustrate the Cu(I)-binding features of the whole series.The titration of the peptides with Cu(I) at pH 7.4 was followed

by UV-Vis spectroscopy. The absorbance stabilizes surprisinglyslowly with the equilibrium reached only after ca. 2 hours. Suchlong equilibration times were not observed before in similarsystems with thiolate containing peptides or pseudopeptides,even when Cu(I) clusters were formed from the very beginning ofthe titration.28,38,39 Titrations using individual samples weretherefore conducted with Cu(I) to peptide concentration ratiosfrom 0.0 to 3.0 in 0.25–0.5 steps to allow for long equilibrationtimes and ensure that the thermodynamic equilibrium isreached. Fig. 1 shows the results of the UV titration of peptide1C with Cu(I) at equilibrium. An intense band, characteristic forthe S� to Cu(I) charge transfer transition (LMCT),70 emerges atl = 263 nm with increasing Cu(I) concentration. The absorbanceincreases linearly up to 2.0 equiv. of Cu(I). The breakpointobserved for 2 equiv. of Cu(I) per peptide is indicative of theformation of polymetallic species with (Cu2P)n overall stoichio-metry as previously observed with 3-Cys-containing tripodalpeptide-like ligands.38,39 As seen before, the intensity of theS� to Cu(I) LMCT is not sensitive to the composition of thecomplex and provides information only about the numberof Cu(I)–thiolate bonds.38,39,71,72 The extinction coefficientse = 6000–7000 M�1 cm�1 per bound Cu(I) reported in Table 1are in good agreement with data reported with other thiolate–Cu(I) complexes.71,73

Cu(I) complex formation was also followed by recordingCD spectra of the individually prepared samples. As a resultof Cu(I) additions, the spectra significantly change comparedto the free ligand and signals appear in the S� to Cu(I) LMCTregion (260–320 nm). However, the molar ellipticities are

Scheme 1 Sequences of the studied peptides. Bonds in the turn motif are indicated in bold and the coordinating cysteine residues are highlighted in red.

Fig. 1 UV spectra of 1C titrated with Cu(I) (equilibration time = 2 hours).The inset shows the increase of the absorbance at 263 nm as a function ofcCu(I)/cpeptide ratio (cpeptide = 30 mM in phosphate buffer, 20 mM, pH = 7.4).

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relatively low ([Y] o 1 � 104 deg cm2 dmol�1) compared tothose obtained with other species involving 3 coordinated Cys([Y] = 4 � 104 – 8 � 104 deg cm2 dmol�1).38,39,71 Moreover, noclear tendency can be observed in the evolution of the spectraupon Cu(I) addition up to 2.0 equivalents, as shown in Fig. S2(ESI†). The mostly affected region falls below 250 nm, where thenegative CD band is attributed to p - p* and possibly over-lapping n - p* transitions, both belonging to the amide bondsof the peptide backbone,74,75 which suggests a conformationalrearrangement upon Cu(I) addition. Since CD spectroscopy ismore sensitive to the structure around Cu(I) than UV, theseresults indicate that the free peptides transform into more thanone Cu(I) species. This assumption is confirmed by ESI-MSmeasurements in ammonium acetate buffer at pH 6.9 (Fig. S3,ESI†). Spectra recorded for samples with the peptides and0.9 equivalents of Cu(I) show the formation of mononuclearcomplexes, CuP, and several polynuclear species, like Cu4P2

and Cu4P3. With the increase of the Cu(I) concentration, furtherspecies of higher nuclearity (Cu8P4, Cu7P3, Cu9P3) are detected.Therefore, the CD and ESI-MS experiments demonstrate thatthe apparently simple evolution of the LMCT bands in the UVtitration is in fact due to the formation of a mixture of manypolynuclear thiolate complexes.

Copper binding affinities were determined in the presenceof BCS as a competitor. BCS forms a well-characterisedCu(BCS)2 complex with Cu(I) according to eqn (1).

Cu+ + 2BCS2� " [Cu(BCS)2]3� log b = 19.8 (1)

This complex has an intense orange colour and a maximalabsorbance at l = 483 nm with an extinction coefficiente = 13 300 M�1 cm�1.51 Solutions of the Cu(I)–peptide complexes(Cu(I) : P ratio = 0.9 : 1) were titrated with BCS and the amount ofCu(I) displaced from the peptide by BCS was quantified based onthe known absorption of the Cu(BCS)2 complex. Since severalCu(I) complexes are formed with the six peptides, the fit of thespectroscopic competition results could not be perfectly imple-mented with a given complex stoichiometry. Hence, the apparentstability constants were calculated considering the formation ofa CuP complex. The logbpH7.4

CuP values are presented in Table 2.

All peptides display high affinity towards Cu(I) in the range typicalof Cu(I) chaperone proteins, like Atx1.22

Hence, Cu(I) binding to the investigated 3-Cys containingpeptides is characterized by a rather complicated speciation atpH = 7.4. Indeed, although the UV profile appears simple with asingle endpoint at 2 equivalents of Cu(I), ESI-MS and CD clearlyevidence several clusters, even with low Cu(I) concentration.This complicated mixture of species observed for the wholeseries of peptides highly contrasts with previous resultsobtained with tripodal pseudopeptides also incorporating threecysteine moieties, which form Cu(I) complexes, resemblingthose formed in metallothioneins.39,71,76 Indeed, the pre-orientation of the three thiolate groups in the tripodal pseudo-peptides induces a well-defined metal binding cavity, stabilizedby a network of hydrogen bonds.39 This structure perfectlycontrols the speciation of the Cu(I) complexes with only twoidentified species, namely the mononuclear complex and thecluster Cu6S9 both with Cu(I) ions in trithiolate environments.39,76

More recently, the highly constrained tetrapeptide Ac-Cys-D-Pro-Pro-Cys-NH2 with a strong turn was shown to form exclusively a Cu4S6

core.77 The larger flexibility of the peptides described here couldbe responsible for the lack of control of the speciation of theCu(I) complexes and ultimately for the formation of a mixture ofpolymetallic species, with quite large stability. The determinedapparent stability constants are similar in the whole peptide series,within the range of experimental errors, indicating that the struc-tural differences have only a minor effect, if any, on the stability ofthe Cu(I) complexes.

Hg(II) complexes

Hg(II) is a metal ion with soft character according to Pearson’stheory,78 and as such, an often used probe of the oxygen andwater sensitive Cu(I). The cyclic peptide P3C has been recentlydemonstrated to form a complex with Hg(II) in a HgS3 coordi-nation environment, which is stable over a large pH range.Besides, the protonation of the mononuclear complex HgP3C

happens at a relatively low pH (pKa value of 4.3) to produce aspecies with HgS2 geometry.40 Hg(II) binding of the new seriesof peptides was therefore studied to reveal whether the beha-viour of P3C with Hg(II) is specific to the structure of this cyclic

Table 1 LMCT band characteristics for Cu(I) and Hg(II) complexes of thestudied peptides

CuP HgP

l (nm) e (M�1 cm�1) l (nm) e (M�1 cm�1)

P3C 259 7500 240 17 50040

270 10 2001C 267 6700 240 13 800

280 73001L 267 6100 240 14 900

280 94002C 267 6200 240 15 000

280 77002L 262 6500 240 14 600

280 77003C 261 6400 240 14 500

280 7800

Table 2 Apparent stability constants, deprotonation constants and someestimated stability data (all in the form of their logarithms) of the Hg(II) andCu(I) complexes. For the experimentally determined values errors on thelast characters are indicated in parentheses

logbpH7.4CuP

a log bpH2.0HgP log bpH7.4

HgPb pKHgL

HgHLc log bHgHL

b logbHgLb

P3C 18.1(1) 27.2(1) 40.9 4.3(1)40 48.7 44.41C 17.3(3) 27.3(1) 40.5 4.8(1) 48.8 44.01L 17.4(2) 27.1(1) 40.0 5.1(1) 48.6 43.52C 17.8(4) 27.2(1) 40.7 4.5(1) 48.7 44.22L 17.8(2) 27.0(1) 40.0 5.0(1) 48.5 43.53C 17.9(4) 27.5(1) 41.0 4.5(1) 49.0 44.5

a The apparent stabilities were calculated by assuming the presence ofonly mononuclear complexes. b Apparent stability constants for pH 7.4and formation constants of the HgHL and HgL species were estimatedfrom the apparent stabilities obtained at pH = 2.0, as described in theESI. c The pK values characterize the HgHL " HgL + H+ process.

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peptide or common for peptide sequences encompassing threeCys residues in a CxCxxC or CxCxC arrangement.

Hg(II) binding at physiological pH. The S� to Hg(II) LMCTbands of mercury–thiolate complexes are indicative of themetal ion coordination number and geometry, in contrast tosimilar bands of Cu(I)–thiolates. The HgS3 coordination modeis characterized by an LMCT band in the wavelength range of240–320 nm,37,79 while the di-coordinate HgS2 structures leavetheir fingerprint only in the higher energy UV region.28,80,81

Accordingly, the Hg(II) binding properties of the peptides werestudied by UV-Vis spectroscopy at pH = 7.4. Titrations wereconducted by gradually adding 0.1 equivalent of Hg(II) to thepeptide solutions. The absorbance values stabilized after Hg(II)additions much faster (in less than 5 minutes) compared to thecomplexation of Cu(I) by the same peptides, which might be aresult of a simpler speciation in the Hg(II)–ligand systems.All peptides showed very similar behaviour to P3C.40 The UVtitration of 1C is presented in Fig. 2 as an example. With theincrease of metal ion concentration, two intense bands emergeat l B 240 and 280 nm with molar absorption coefficients(Table 1) similar to those previously reported for HgS3 typecomplexes.36,37,80 Absorbances increase linearly up to 1 : 1Hg(II)–peptide ratios, which reflects the formation of a singlemononuclear complex, where Hg(II) is very likely coordinated bythe three cysteine thiolates of the peptides. In the presence ofHg(II) in excess, a decrease of the absorbances at the selectedwavelength values can be observed, indicating that the mono-nuclear complex transforms into polymetallic species with Hg(II)ions coordinated by only two cysteine thiolate residues.28,80,81

Besides, the second endpoint observed at 1.5 equiv. of Hg(II)strongly suggests the formation of Hg3P2 complexes.

ESI-MS experiments performed in ammonium acetate bufferat pH 6.9 support well the UV spectroscopic results (Fig. 3).The spectrum of 1C with 1.0 equivalent of Hg(II) clearly showsthe presence of the mononuclear HgP complex in the forms of a[Hg1C]2+ and a [Hg1C–H]+ ion with m/z = 576.8 and 1152.3,respectively. The spectrum recorded with 1.5 equivalents ofHg(II) indicates the formation of new species. The majordetected complex is Hg3P2 in the forms of [Hg3(1C)2–4H + Na]3+

and [Hg3(1C)2–4H]2+ with m/z = 841.7 and 1250.8, respectively.This species is compatible with the HgS2 coordination environ-ment of the metal ions. When 2.0 equivalents of Hg(II) are added,Hg3P2 and Hg2P ([Hg2(1C)–2H]2+; m/z = 675.8) can be detected insimilar amounts. This mixture of complexes may explain theslight increase of the absorbance above 1.5 equivalents of Hg(II)per peptide.

All the peptides showed the formation of Hg(II) complexeswith a simple speciation as described recently with the P3C

cyclic decapeptide.40 The mononuclear complexes with featurescharacterizing a trithiolate coordination are formed at physio-logical pH, as also extensively reported for the Hg(II) complexesof triple coiled-coil peptides.34,35 Contrary to the latter systems,which define a highly protected hydrophobic metal-bindingpocket, the flexible structure of the cyclic and linear peptidesmakes the transformation of the trithiolate coordination intothe preferred dithiolate coordination possible in excess of Hg(II)in Hg3P2 complexes.

Protonation of the HgL complex. The absorbance at 280 nm,characteristic of the HgS3 geometry in the Hg(II) peptidecomplexes, decreases significantly with decreasing pH, whichindicates the protonation of one cysteine to afford a linear HgS2

complex at low pH, as observed previously with P3C.40 Fig. 4shows the pH titration for Hg1C as an example. The spectroscopicdata were satisfactorily fitted with one pKa value according to thefollowing equation:

HgHLÐ HgLþH KHgLHgHL ¼

½HgL�½H�½HgHL� (2)

where [HgHL] and [HgL] denote the equilibrium concentrationsof the mononuclear complexes including the peptide in differentprotonation states.

Fig. 2 UV titration of 1C with Hg(II) in phosphate buffer (20 mM, pH = 7.4).The inset shows the increase of the absorbance at 240 and 280 nm as afunction of cHg(II)/cpeptide ratio (cpeptide = 30 mM).

Fig. 3 (+)ESI-MS spectra registered for 1C with different equivalents ofHg(II) in ammonium acetate buffer (20 mM, pH = 6.9). The comparison ofthe experimental and calculated isotopic envelop of the detected speciesis also presented. Asterisks mark the sodium adduct of the correspondingspecies. The notation 1C refers here to the neutral free peptide.

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The pKa of the Hg(II)–peptide complexes (see Table 2) followthe order of P3C o 3C E 2C o 1C o 2L E 1L. However, thedifferences observed between the pKa values in the series ofpeptides (maximum difference = 0.8) are quite small, whichreflects the weak influence of the pattern of the three Cysresidues (CxCxC or CxCxxC) and the separation of the metalbinding fragment from the turn motif. P3C incorporating theCxCxxC sequence forms the HgS3 coordination mode at aslightly lower pH than the other cyclic peptides. A largerseparation from the PG-turn also seems to be favourable forthe formation of the tris-thiolate complex in 3C. The highestpKa values are seen for the two linear peptides, which may be aconsequence of their larger flexibility and a more significantreorganization necessary for the coordination of the third Cyssidechain. It is rather interesting that even the latter data(Table 2) are 1.7–1.8 log units lower than the pKa observed forthe Hg(II)-complex of the tris-cysteine functionalized tripodalpseudopeptide ligand with amidated carboxyl groups,37 orof the three-stranded coiled coils (pKa values of 8.6 and 7.6for sites d and a, respectively).34 This might be related tothe presumably very different water-accessibility of the thiolgroups, as hinted recently.40

Stabilities of the Hg(II) complexes. Due to the high thio-philicity of Hg(II), the determination of the stability of thecomplexes is rather challenging. Competition titrations withiodide ions were performed to measure the apparent stabilityconstants at pH = 2.0. To the best of our knowledge, this is thefirst time that the well-characterized Hg(II)-complexing abilityof iodide has been utilized for the determination of the stabilityof Hg(II)–thiolate complexes, which is probably due to thepresence of several iodo complexes. According to the observedpH-dependent transformation of the Hg(II)–peptide complexes,the apparent stabilities determined at pH = 2.0 mostly correspondto the species with a HgS2 structure. Hg(II) has been shown toform iodo complexes in four consecutive steps characterized bylarge formation constants.56 In spite of these large iodo complexstabilities, a high concentration of I� (1500–2000 equiv.) was

necessary to withdraw Hg(II) from the peptide complexes whichrequired the use of a background electrolyte (I = 0.1 M NaClO4) tolessen the change in the ionic strength during the titrations.

Therefore, the published stabilities of the iodo complexeswere recalculated by applying the SIT model to the conditionsof the experiments,57,58 leading to the following formationconstants: log b[HgI]+ = 13.05, log b[HgI2] = 24.09, log b[HgI3]� =27.84, logb[HgI4]2

� = 29.91. The result of the titration ofHg(II) with KI and the obtained molar spectra of the formingHg(II)–I� complexes can be seen in Fig. 5A and Fig. S4 (ESI†),respectively.

Samples of the peptides containing 1.0 equivalent of Hg(II)were titrated with I� and the recorded spectra are presented inFig. 5B. The first phase of the titrations, i.e. up to the presenceof 10 equivalents of I�, does not reflect considerable changes inthe recorded spectra. Further addition of I� ions results in theappearance of new bands characteristic for the [HgI3]� and[HgI4]2� species (compare to the spectra in Fig. S4, ESI†).A complete displacement of Hg(II) from the peptide complexesis achieved at ca. 2000 equiv. of I�.

The obtained spectra were fitted by SPECFIT by fixing thelog b values and the molar spectra of the Hg(II)–I� complexes.The best fits were obtained when the formation of a mixedligand complex, HgPI, was also included in the models, besidesthe HgP species (inset in Fig. 5B). The appearance of suchspecies is probably a consequence of the flexibility of thepeptide structures. The HgPI complexes are present only in

Fig. 4 pH titration of 1C with 1.0 equivalent of Hg(II). The dashed linerepresents the spectrum of free 1C at pH = 2. The inset shows theevolution of the absorbance as a function of pH at 280 nm. Symbolsrepresent the experimental points and the line is the fitted absorbanceobtained by SPECFIT.

Fig. 5 UV spectra recorded in the titration of (A) Hg(II) and (B) Hg1C withI�. pH = 2.0, cHg(II) = 30 mM (A), cHg(II) = cpeptide = 30 mM (B). The insets showthe evolution of the absorbance at 322 nm (K) and 280 nm (J). Symbolsrepresent the experimental data, and solid lines represent the absorbancescalculated by SPECFIT.

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the beginning part of the titrations in a ca. 20% relativeproportion, except for HgP3C where a somewhat larger fractionof HgPI could be observed. The formation of the mixedligand complexes, according to the HgP + I " HgPI equation, ischaracterized by stabilities falling in the range of log K B 1.5–2.5.The apparent stability constants determined for the HgPcomplexes (Table 2) indicate rather similar affinities of Hg(II)to all peptides. Considering that at pH = 2.0 Hg(II) is coordi-nated only by two thiolate units, it is a plausible assumptionthat the preorientation of the donor groups has only a modestinfluence on the stabilities and the high thiophilicity of Hg(II)easily governs the formation of the favoured HgS2 structures.Formation constants for the different forms of the HgP species,i.e. for HgHL and HgL (where L denotes the fully deprotonatedpeptide), and apparent stabilities for pH = 7.4, were alsoestimated from the relevant conditional stability constants(log bpH2.0

HgP ) by applying the pKa values of the HgHL " HgL +H processes (Table 2) and the pKa values obtained for one of thefree ligands, pKHL

a = 9.26(1); pKH2La ¼ 8:56ð1Þ; pKH3L

a ¼ 7:67ð1Þ(see the Experimental part and the ESI†). The stability con-stants estimated for the Hg(II)–trithiolate HgL complexes spanover a small range (maximum difference, Dlog bHgP = 1) demon-strating a weak influence of the Cys-sidechain orientations inthe chosen sequences (Table 2). Nevertheless, the two linearpeptides display a slightly weaker affinity suggesting the needfor a more pronounced rearrangement of the Cys sidechains.It is noteworthy to compare these data to the stability constantsof Hg(II)–bisthiolate complexes of highly constrained bis-thiolligands. Our peptides, indeed, display very similar Hg(II)-bindingaffinities to that of the well-known soft metal ion chelator2,3-dimercaptopropan-1-ol (BAL) (logbHgL = 44.8),82 which formsa highly stable 5-membered chelate ring around the metal ion.Comparison of our data to the stability of the HgL species of thetetrapeptide CDPPC (logbHgL = 40.0)81 clearly indicates that thestructure of our peptides is prone to easily rearrange to a suitableform for the tridentate coordination of Hg(II) and thus the largernumber of Hg(II)–thiolate bonds is revealed by higher affinities.

Modelling of the trithiolate HgL complexes

The six peptides were modelled in the apo and Hg(II)-boundform with a HgS3 trigonal coordination (see Experimentalsection for methods). Several independent simulations wererun: first, for the Hg(II)-bound peptides three different simula-tions were run varying the orders of the cysteine sulfur atomsdefining the Hg–S–S–S improper angle; then, for each bestenergy conformation in each system, new simulations for therelated systems were run starting from there. The energy valuesreported in Table S2 (ESI†) refer to the simulation leading tothe minimum average total energy for each peptide (overall simulations). Binding of Hg(II) to linear or cyclic peptidesalways results in stabilization meaning that the peptide struc-ture organizes the 3 cysteine residues in an environmentappropriate for HgS3 coordination. The stabilization expressedas DE = E(HgP) � E(P) is remarkably similar for all four cyclicpeptides (between�11 and�12.8 kcal mol�1 from Table S2, ESI†)

with a relative error (DE/E) less than 1%. This stabilization variesfrom �13.1 to �9.5 kcal mol�1 for the two linear peptides;however, this slightly larger difference may be due to anincomplete conformational search of the highly flexible linearapo peptides. Hence, the calculated stabilization energies, DE,are very similar in the series of peptides, which is in agreementwith the comparable values of the stability constants of the HgLcomplexes determined experimentally (Table 2). Models ofenergy minimized structures of the Hg(II)-bound peptides,starting from the frame with the lowest potential energy duringthe dynamics simulation leading to minimum average totalenergy, are shown in Fig. 6 and Fig. S5 (ESI†) for cyclic andlinear peptides, respectively. The three cysteine binding chainsare well-disposed to afford the trithiolate coordination of themercury ion in the six structures. In all the complexes, thepositively charged sidechain of arginine (charge +1) is cappingthe HgS3, negatively charged binding site (charge �1), providingstabilizing electrostatic interactions.

Conclusions

Model peptides containing cysteine-rich sequences found inmetallothioneins were studied for their metal-binding proper-ties in relation to metal detoxification mechanisms. The twosoft ions Cu(I) and Hg(II) were selected since they exhibit thelargest affinities for these small detoxification proteins amongendogenous and toxic metal ions, respectively. Three cysteineresidues were introduced in CxCxxC and CxCxC motifsin different positions within the sequence, in linear and cyclicderivatives. Overall, the six peptides display rather similarbehaviour, which evidences minor contributions of the positionof the three cysteine residues or cyclisation to the formationand stability of the Hg(II) and Cu(I) complexes.

Cu(I) binding to the series of peptides at physiological pHrevealed to be rather complicated, with the formation of amixture of polymetallic species. In contrast, cysteine-rich highlystructured peptides77 or peptide-like ligands39,76 are able to

Fig. 6 Energy minimized structures of the 4 cyclic peptides in theirHg(II)-bound form (oriented with respect to the position of backboneatom coordinates of residues 1 to 10).

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control the formation of well-defined Cu(I) complexes. Conse-quently, the complicated Cu(I)-complex speciation of the seriesof peptides, reported in this paper, has been assigned to theirsignificantly larger flexibility. However, despite the formationof many polymetallic species, large affinity is achieved for thesoft Cu(I) cation at physiological pH (1017–1018).

The binding of Hg(II), another soft metal ion often used as aprobe for the oxygen and water sensitive Cu(I), demonstratesthat the complexity of the Cu(I) speciation is due to the peculiarbehaviour of Cu(I)–thiolate complexes in water and not to thecysteine-rich sequences chosen for the peptides. Interestingly,the HgS3 coordination mode is stable over a large pH-range forall studied peptide complexes. Indeed, the protonation of thecomplex to give the HgS2 linear coordination is observed withpKa values ranging from 4.3 to 5.1, making the trithiolatecoordination the major binding mode at physiological pHwhatever the peptide sequence. The stabilities of the Hg(II)complexes (1040–1041 at pH 7.4) are of the same order ofmagnitude as those reported for high affinity sulphur chelatingagents such as BAL.82 The large stability constants togetherwith the low pKa values and simulated structures clearly indicatethat all the peptide sequences studied in this paper are adaptedfor an efficient trithiolate coordination of the thiophilic cationHg(II).

Importantly, the striking differences observed in the coordi-nation of Hg(II) and Cu(I) with the series of peptides indicatedifferent molecular mechanisms involved in their bindingto detoxification proteins. The sulphur-rich peptides studiedhere show more than 20 orders of magnitude larger affinity atpH 7.4 for Hg(II) (log bpH7.4

HgP E 41) than for Cu(I) (log bpH7.4CuP E

18), due to the significantly softer character of Hg(II). Mostimportantly, Hg(II) forms well-defined complexes, whereasCu(I)-coordination leads to mixtures of polymetallic species.This demonstrates the peculiar behaviour of Cu(I) thiolatecomplexes in water. Only highly constrained peptide sequencesare able to promote the formation of well-defined Cu(I) complexes.The peptides studied here are probably too flexible to achieve sucha control for Cu(I). Hence, the use of Hg(II) as a probe for Cu(I)coordination with sulphur-rich peptides or proteins in physio-logical conditions is demonstrated here to be not fully appropriate.

Conflicts of interest

There are no conflicts to declare.

Acknowledgements

This research was supported by the Labex ARCANE (Grant ANR-11-LABX-0003-01), the ‘‘Fondation pour la Recherche Medicale’’(grant DCM20111223043), the Hungarian National Research,Development and Innovation Office-NKFIH through projectGINOP-2.3.2-15-2016-00038 and grant no. K_16/120130. EditMesterhazy kindly acknowledges the financial support ofCampus France.

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